Characterization, efficient transformation and regeneration of Chirita pumila (Gesneriaceae), a...
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ORIGINAL PAPER
Characterization, efficient transformation and regenerationof Chirita pumila (Gesneriaceae), a potential evo-devo model plant
Bo-Ling Liu • Xia Yang • Jing Liu •
Yang Dong • Yin-Zheng Wang
Received: 12 December 2013 / Accepted: 4 April 2014
� Springer Science+Business Media Dordrecht 2014
Abstract An efficient transformation and regeneration
system is essential for functional investigation of devel-
opmental genes and related elements in the field of evo-
lutionary developmental biology (evo-devo). Chirita
pumila D. Don belongs to the Gesneriaceae family, one of
the most basal groups in Lamiales sensu lato, and possesses
many tractable biological features including annual habit,
small plant size, short generation time, abundant offspring
and low chromosome number. In addition, C. pumila has
cleistogamous flowers with potential cross-pollination, a
special phenomenon first reported herein in Gesneriaceae.
Parameters affecting shoot induction and genetic transfor-
mation have been evaluated, including plant growth regu-
lators, temperature, antibiotic concentration, pre- and co-
culture duration, Agrobacterium cell density and infection
time. Polymerase chain reaction and b-Glucuronidase
(GUS) activity assays of T0 and T1 plants show that the
GUS gene has been introduced into the host with stable and
universal expression. The applicability of the transforma-
tion system in gene function investigation is further con-
firmed by transforming a GsNST1B gene from Glycine
soja. This transformation system provides a valuable
platform for deep function analyses of related genes and
elements for a wide range of evo-devo studies, especially
in the field of floral evolution, which would develop its
potential of being a model organism in Lamiales s. l.
Keywords Chirita pumila � Cleistogamy � Evo-devo �Genetic transformation � Gesneriaceae
Abbreviations
BA 6-Benzylaminopurine
GFP Green fluorescent protein
GUS b-Glucuronidase
HPTII Hygromycin phosphotransferase gene
MS Murashige and Skoog medium
NAA 1-Naphthaleneacetic acid
PCR Polymerase chain reaction
RT-PCR Reverse transcription-PCR
Introduction
Only after the rise of evolutionary developmental biology
(evo-devo) has the integration of developmental processes
and genetic and evolutionary biology at the molecular level
allowed the analysis of how developmental processes can
result in morphological evolution (Breuker et al. 2006;
Kellogg 2006; Muller 2007; Carroll 2008; Kopp 2009; de
Bruijn et al. 2012). Over the past two decades, evo-devo as
an emerging biological discipline has made considerable
achievements in discovering extensive similarities in gene
regulation among distantly related species with dramati-
cally different body plans in both animals and plants
relying on rapid technical advancements in gene clone and
Bo-Ling Liu and Xia Yang have contributed equally to this work.
Electronic supplementary material The online version of thisarticle (doi:10.1007/s11240-014-0488-2) contains supplementarymaterial, which is available to authorized users.
B.-L. Liu � X. Yang � J. Liu � Y. Dong � Y.-Z. Wang (&)
State Key Laboratory of Systematic and Evolutionary Botany,
Institute of Botany, Chinese Academy of Sciences,
20 Nanxincun, Xiangshan, Beijing 100093, China
e-mail: [email protected]
B.-L. Liu
Qufu Normal University, Qufu, Shandong, China
123
Plant Cell Tiss Organ Cult
DOI 10.1007/s11240-014-0488-2
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expression. Examples include Hox genes in establishing the
anterior–posterior axis in bilaterian animals, MADS-box
genes in patterning floral organ identities and CYCLOIDEA
(CYC)-like TCP genes in determining floral zygomorphy
(Breuker et al. 2006; Kellogg 2006; Muller 2007; Carroll
2008; Kopp 2009; de Bruijn et al. 2012). These studies
have provided remarkable insights into the evolutionary
conservation of developmental programs and the mecha-
nisms underlying modification of developmental processes
that generate morphological novelties (Stern 2000; Pruitt
et al. 2003; Breuker et al. 2006). Currently, most evo-devo
studies in plants, especially outside model organisms,
depend on global DNA sequence analyses and correlative
analyses of candidate gene expression to corresponding
morphologies rather than gene function investigation. Even
though gene expression studies are sufficient ways to
screen genes for putative regulatory changes, they are
association rather than causality analyses (Baguna and
Garcia-Fernandez 2003; Kellogg 2004). Therefore, it is
essential and critically important to conduct comparative
functional studies to demonstrate that such regulatory
changes are actually responsible for phenotypic variations
and to gain an integrated view of the role of development
in evolution (Irish and Benfey 2004; Breuker et al. 2006).
As functional analyses become widespread in evo-devo
studies, researchers usually transfer target genes into a
distantly related classical model organism to test their
function because of the difficulty in carrying out such
experiments in native systems (Irish and Benfey 2004).
However, these gene transfers are not always efficient to
test the genes’ function or may not reflect their actual
function in native contexts (Irish and Benfey 2004).
Therefore, evo-devo biologists have increasingly recog-
nized the limitation of the classical model organisms and
the urgency to develop new model organisms to efficiently
investigate the genes’ actual function in specific morpho-
logical novelties (Mandoli and Olmstead 2000; Irish and
Benfey 2004; Jeffery 2008).
In angiosperms, one key innovation is the occurrence of
the flower with subsequent remarkable diversification upon
wide modifications of the genetic programs controlling
floral organ identity, floral symmetry and reproduction
system (Dilcher 2000; Kramer 2007). Currently, the focus
of plant evo-devo studies is mainly on the evolution and
diversity of ABCE model beneath floral organ identity and
gene network underlying floral symmetry first identified
and elaborately studied in classical model species Arabi-
dopsis and Antirrhinum (Irish and Benfey 2004; Kramer
2007). New evo-devo model organisms would yield new
insights into the origin of major floral evolutionary nov-
elties in particular lineage histories that could not be tar-
geted by classical model organisms. The Gesneriaceae
family is one of the most basal groups in Lamiales sensu
lato (Endress 1998; Cubas 2004; Wortley et al. 2005; http://
www.mobot.org/MOBOT/Research/APweb/welcome.html),
a major angiosperm clade predominant with zygomorphic
flowers that are believed ancestral in this order (Donoghue
et al. 1998; Cubas 2004). Therefore, Gesneriaceae locates at
a phylogenetic node of floral evolution in angiosperms. As a
member of Gesneriaceae, Chirita pumila D. Don is a
promising candidate of model species for evo-devo studies
in floral evolution because it shares a series of biological
features with classical model plants, such as annual habit,
diploid with the lowest chromosome number (2n = 8) in
Gesneriaceae (Ratter 1963; Li and Wang 2004; this study),
and cleistogamy with potential cross-fertilization (see
results in this study). In addition, the whole genome
sequencing project is carrying out (Yi-Kun He, personal
communication). These unique biological features give
C. pumila a great advantage in tractability for laboratory
experiments over other Gesneriaceae species that are usually
perennial and polyploidy with cross-pollination, including
the two famous ornamental plants Saintpaulia ionantha and
Sinningia speciosa and the physiological model plant Ra-
monda myconi successful in Agrobacterium-mediated
genetic transformation (Mercuri et al. 2000; Kushikawa
et al. 2001; Toth et al. 2006; Zhang et al. 2008).
Our laboratory has conducted a series of evo-devo
studies relating to the evolution of floral symmetry in
Gesneriaceae (Du and Wang 2008; Gao et al. 2008; Zhou
et al. 2008; Song et al. 2009; Pang et al. 2010; Yang et al.
2010; Liu et al. 2014) and the molecular mechanism
underlying the repeated origins of floral zygomorphy in
angiosperms (Yang et al. 2012). Recently, deep functional
analyses of related gene networks in floral symmetry and
floral organ identity are carrying out in C. pumila and its
relatives (our unpublished results). Herein, we report an
efficient Agrobacterium-mediated transformation and
regeneration system developed in C. pumila by evaluating
several factors affecting shoot induction and genetic
transformation and validating its applicability in gene
function investigation using the GsNST1B gene functioning
in secondary wall biosynthesis in Glycine soja. This
transformation system would have wide applications in the
field of evo-devo studies.
Materials and methods
Plant material and culture conditions
The C. pumila plants, collected from Hekou County,
Yunnan, China (Wang, HK01), were grown in 8 cm pots
containing the mixture of vermiculite and Pindstrup
substrate (Pindstrup) (1:2) in culture room. The growth
conditions were: 26 �C, a 10/14 light/dark photoperiod
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under cool-white fluorescent light of 100 lmol m-2 s-1
and 50–70 % of relative humidity. Voucher specimens
were deposited in the Herbarium, Institute of Botany,
Chinese Academy of Sciences.
Agrobacterium strain and plasmids
Agrobacterium tumefaciens strain LBA4404 harboring the
binary vector pCAMBIA1301 was used in parameter eval-
uation experiments and b-Glucuronidase (GUS) activity
assay. The vector carries the hygromycin phosphotransfer-
ase (HPTII) gene for transformant selection on hygromycin
and the GUS reporter gene that is interrupted by an intron
(Fig. 1a). The p35S::GsNST1B plasmid was constructed as
described (Dong et al. 2013). Briefly, the full-length coding
sequence of GsNST1B gene (or GsSHAT1-5; Dong et al.
2013, 2014) was amplified (50-GGAAGATCTGCCGGA
AAACATGAG-30 and 50-GGACTAGTCTACACTG ACG
TGTTGGAC-30), digested with BglII and SpeI, and inserted
into the binary vector pCAMBIA1302 that contains the
HPTII gene and the green fluorescent protein (GFP) gene
(Fig. 1b). The resultant construct was introduced into
Agrobacterium LBA4404 by electroporation (Eppendorf).
Pollen germination assay and aniline blue staining
of pollen tubes
Pollen germination experiment was performed according to
Mori et al. (2006). Briefly, pollen was randomly collected
from six different flowers close to anthesis and dispersed into
sterilized water. 15 ll of the suspension was carefully
flattened onto the culture medium (containing 150 g l-1
sucrose, 40 mg l-1 boric acid, 20 mg l-1 calcium chloride,
6 g l-1 agar), cultured at 28 �C in the dark for 2–8 h and
examined using a Zeiss Axio Imager A1 M Microscope
(Zeiss).
To perform aniline blue staining, flowers close to
anthesis were either collected directly or bagged in Cel-
lophane for further 2 days. The pistils were dissected and
stained with aniline blue according to Jiang et al. (2005).
Briefly, the pistils were fixed in ethanol: chloroform: acetic
acid (6:3:1) for 24 h, softened in 8 M NaOH for 6 h and
washed three times with 0.1 M K2HPO4-KOH buffer (pH
7.5). Then, the pistils were stained in 0.1 % aniline blue
solution (pH 11) in the dark for 4 h and observed with a
Leica TCS SP5 Fluorescence Microscope (Leica).
Karyotype analysis and measurement of the genome
size
Root tips were pretreated with a mixture of 2 mM
8-hydroxyquinoline and 0.1 % colchicine (1:1) at 20 �C for
4 h, and fixed in Carnoy’s I (100 % ethanol and glacial
acetic acid, 3:1) at 5 �C for 1 h. The fixed materials were
macerated in 1 M HCl at 60 �C for 1 min, stained with
carbol fuchsin, squashed and photographed under a
microscope. The length of long and short arms of each
metaphase chromosome was measured, and the chromo-
some arm ratio was estimated by the length of long arm/the
length of short arm. The relative length was calculated by
the length of individual chromosome/the length of all
chromosomes 3100 %.
Fig. 1 Map of the binary vector
pCAMBIA1301 and the
p35S::GsNST1B construct.
a T-DNA region of the
pCAMBIA1301 vector. b The
binary vector pCAMBIA1302
and the constructed
p35S::GsNST1B plasmid
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Flow cytometry was used to measure the genome size of
C. pumila according to Dolezel et al. (2007). Rice (Oryza
sativa L. var. Nipponbare) was used as an internal standard.
Briefly, young leaves of the sample and reference standard
were chopped quickly with a sharp razor blade in a plastic
Petri dish containing 1 ml ice-cold Galbraith’s buffer
(45 mM MgCl2, 20 mM MOPS, 30 mM sodium citrate,
0.1 % Triton X-100, pH 7.0) (Galbraith et al. 1983). The
resultant homogenate was filtered through a 25 lm nylon
mesh to remove large debris, and incubated in staining
solution containing 50 g l-1 propidium iodide (Sigma-
Aldrich) and 50 g l-1 RNaseA (TaKaRa) on ice in the dark
for 20 min. The relative nuclear DNA fluorescence inten-
sity was measured using a MoFlo� High-performance Cell
Sorter (Beckman). Three C. pumila plants were analyzed
with three replicates each.
Seed germination, shoot induction and antibiotic
sensitivity experiments
Chirita pumila plants were grown to flowering stage and
seeds were harvested. The seeds were surface-sterilized in
70 % ethanol for 1 min, rinsed with sterile water once,
disinfected with 2.5 % sodium hypochlorite for 3–5 min
and finally rinsed five times with sterile water. The steril-
ized seeds were germinated on Murashige and Skoog (MS)
medium (Murashige and Skoog 1962) in a growth chamber
at 26 �C under a photoperiod of 10/14 h light/dark
(100 lmol m-2 s-1). About 2-month-old plantlets were
used for preparing leaf explants.
To determine optimal concentration of growth regula-
tors for shoot induction, fresh leaf explants were cultured
on MS medium containing different concentrations of
6-benzylaminopurine (BA) and a-naphthalene acetic acid
(NAA) (see Table 1) at 26 �C. To evaluate the effect of
temperature on shoot induction, fresh leaf explants were
cultured on MS medium containing 0.5 mg l-1 BA and
0.1 mg l-1 NAA (based on the result of growth regulator
experiment; see Table 1) at 22, 24, 26 or 28 �C (see Sup-
plementary Table 4). To evaluate whether hygromycin is
appropriate for selecting transformants, fresh leaf explants
were incubated on MS medium containing 0.5 mg l-1 BA,
0.1 mg l-1 NAA and different concentrations of hygro-
mycin (0, 5, 10, 15, 20 and 30 mg l-1; see Supplementary
Table 5) at 26 �C. For each experiment, the explants were
always maintained on the same medium without sub-
culture, and the shoot induction rate was calculated
4 weeks later. Data, presented as mean ± SD, were cal-
culated from three independent experiments with about 40
leaf explants each. Means of induction efficiencies were
compared for level of significance (P \ 0.05) using a
Fisher’s Least Significant Difference (LSD) test.
Evaluation of parameters affecting shoot induction rate
in transformation experiments
Four parameters were successively evaluated, including co-
culture time, pre-culture duration, Agrobacterium cell
density and infection time. In each experiment, only one
factor was changed with other fixed. The following is a
general protocol for these experiments. Fresh leaf explants
were pre-cultured on MS medium containing 0.5 mg l-1
BA and 0.1 mg l-1 NAA for 0, 1, 2, 3 or 4 days. Agro-
bacterium LBA4404 cells (harboring pCAMBIA1301)
cultured in YEB medium (containing 100 mg l-1 strepto-
mycin, 50 mg l-1 rifampicin and 50 mg l-1 kanamycin) at
28 �C overnight were inoculated to fresh YEB medium and
grown to OD600 = 0.2, 0.4, 0.6, 0.8 or 1.0. The cells were
harvested by centrifugation at 5,000 rpm for 10 min, rinsed
with MS liquid once, and resuspended in MS liquid con-
taining 150 mg l-1 acetosyringone. The harvested cells
were used to inoculate pre-cultured explants for 10, 20, 30,
40 or 50 min. After being briefly blot-dried with sterile
filter papers, the explants were incubated on the co-culture
medium containing 0.5 mg l-1 BA, 0.1 mg l-1 NAA and
150 mg l-1 acetosyringone at 26 �C in the dark for 1, 2, 3,
4 or 5 days, and then transferred to the shoot induction
medium containing 0.5 mg l-1 BA, 0.1 mg l-1 NAA,
Table 1 Effects of different concentrations of BA and NAA on shoot
induction rate of C. pumila leaf explants
BA
(mg l-1)
NAA
(mg l-1)
Shoot induction
rate (%)
Root induction
rate (%)
0 0 67.8 ± 10.0 0
1 1 72.7 ± 7.3 42.5 ± 11.5
1 0.5 79.9 ± 5.1 15.0 ± 0.6
1 0.2 84.8 ± 8.0 0
1 0.1 87.2 ± 11.8 0
0.5 1 87.6 ± 4.2 92.5 ± 0.3
0.5 0.5 82.2 ± 9.2 42.3 ± 6.7
0.5 0.2 92.5 ± 0.3 0
0.5 0.1 97.6 – 4.1 0
0.2 1 65.4 ± 13.9 87.4 ± 4.8
0.2 0.5 60.1 ± 8.1 32.2 ± 10.0
0.2 0.2 85.0 ± 0.6 5.0 ± 4.3
0.2 0.1 82.4 ± 4.8 0
0.1 1 89.7 ± 11.8 95.1 ± 4.3
0.1 0.5 59.9 ± 12.2 35.0 ± 3.9
0.1 0.2 89.9 ± 4.6 7.5 ± 0.3
0.1 0.1 74.9 ± 5.0 0
Data are presented as mean ± SD collected from three independent
experiments, including a total of 40 leaf explants per replicate
The optimal concentration of BA and NAA for the shoot induction is
highlighted by bold letters
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20 mg l-1 hygromycin and 300 mg l-1 carbenicillin. The
hygromycin-resistant shoot induction rate was evaluated
4 weeks later. Data (mean ± SD) were calculated from
three independent experiments with about 40 leaf explants
each. Means of induction efficiencies were compared for
level of significance (P \ 0.05) using a Fisher’s LSD test.
GUS activity assay
One-day pre-cultured leaf explants were inoculated with
Agrobacterium LBA4404 (harboring pCAMBIA1301) for
20 min, cultured on the co-culture medium in the dark for
2 days and then transferred to the shoot induction medium.
Hygromycin-resistant shoots of about 0.5 cm in length were
excised, transferred to fresh MS medium (without growth
regulator and antibiotic), and finally transferred to pots and
grown in culture room (the growth conditions were the same
as described above). Genomic DNA was isolated from the
leaves of putative T0 plants using the Rapid Plant DNA
Extraction Kit (Tiangen), and PCR was conducted to in-
dentify positive transgenic plants using primers spanning the
35S promoter and the GUS gene (50-GTGAGCGGATAACA
ATTTCAC-30 and 50-CGAGTCGTC GGTTCTGTAAC-30).PCR conditions were: 94 �C 3 min, 30 cycles of 94 �C 30 s,
60 �C 30 s and 72 �C 60 s, and 72 �C 10 min. Plasmid and
wild-type plants were used as positive and negative controls,
respectively. GUS staining was conducted as described
(Jefferson et al. 1987). Briefly, leaves and stems of three
independent T0 transgenic plants were incubated in GUS
staining solution (50 mM sodium phosphate buffer,
0.05 mM potassium ferricyanide, 0.05 % Triton X-100,
2 mM X-Gluc, pH 7.0) at 37 �C overnight and observed
under a microscope. Wild-type plants were served as nega-
tive controls to exclude the possibility of endogenous GUS
expression. To validate whether the transformed GUS gene
could be inherited, T1 progenies of one T0 plant were dis-
infected and cultured on MS selection medium containing
25 mg l-1 of hygromycin. About 4 weeks later, the segre-
gation ratio was calculated by counting the number of ger-
minated and well developed seedlings and the number of
germinated but withered seedlings. The GUS activity assays
of T1 progenies were as described above.
Expression and histochemical analyses of GsNST1B
gene in transgenic plants
Agrobacterium LBA4404 harboring the p35S::GsNST1B
plasmid was used to infect C. pumila leaf explants. Positive
transformants were confirmed by PCR followed by DNA
sequencing to exclude the possible amplification of
endogenous NST1B-like genes. RT-PCR was conducted to
measure GsNST1B expression in transgenic leaves using
specific primers (50-CTGGCCGCGACAAAGTCATC-30
and 50-CTTCTTCCTGAGCAGCATCCG-30; Dong et al.
2013) under the following conditions: 94 �C 3 min, 30
cycles of 94 �C 30 s, 56 �C 30 s and 72 �C 30 s, and 72 �C
10 min. RT-PCR products were sequenced. As a reference
gene, CpACTIN was amplified with 26 cycles using specific
primers (50-AGTTATCACCATTGCC GCCGAGAGG-30
and 50-GCAATGCCAGGGAACATAGTCGACC-30). RT-
PCR products were visualized on a 1.5 % agarose gel.
The ectopic deposition of lignin was examined as described
(Dong et al. 2013). Briefly, transgenic leaves were fixed,
embedded in Paraplast Plus (Sigma-Aldrich) and stained
with 0.2 % toluidine blue solution. The autofluorescence of
secondary cell walls was detected using a fluorescence
microscope (Zeiss). Three transgenic plants were examined
with wild-type ones served as negative controls.
Results
Biological characteristics of C. pumila plants
Chirita pumila D. Don, an annual herb with erect stems of
6–46 cm in height, extensively distributes in Southwest
China, North India, Vietnam, Nepal, Sikkim, Bhutan, Myan-
mar and Thailand (Wang et al. 1998; Li and Wang 2004). Its
typical characteristics include purple-spotted oval leaves and
large purplish zygomorphic flowers (Fig. 2a). The capsule is
6–12 cm in length (Fig. 2a, b) that can yield abundant tiny
Fig. 2 Morphology of C. pumila. a A C. pumila plant with typical
zygomorphic flowers. Bar, 2 cm. b The capsules of C. pumila. Bar,
2 cm. c The capsule and seeds of C. pumila. Bar, 1 cm. d The
enlarged view of c. Bar, 0.1 cm
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spindly seeds (over 1,000 seeds per capsule based on a rough
estimate) (Fig. 2c, d). In addition, C. pumila has a short gen-
eration time (about 5 months from seed to seed).
Self-compatibility is critical to a genetic transformation
system (Bliss et al. 2013). We have noticed that C. pumila
flowers always autonomously bear fruits without any source
of pollinators in culture room (Fig. 2b). Here, we need to
confirm whether and when the C. pumila flowers are self-
fertile through a series of experiments. We first dissected
longitudinally the flowers just close to anthesis to examine
whether the sexual organs are mature (Fig. 3a, b). Within the
closed corolla tube, the style is held in position pressed
against the upper inner surface of the tube with the bila-
mellar stigma curved downward at the tip (Fig. 3b, c). The
filaments of two stamens strongly geniculate at the mid-
point and lift the two face-to-face cohered anthers above the
stigma and pressing against the style (Fig. 3b, c). There is a
great amount of pollen released from anthers and fallen on
the lower inner surface of the corolla tube (Fig. 3b, c). The
results of in vitro pollen germination experiments further
showed that nearly 100 % of pollen grains started to ger-
minate after incubating on the culture medium at 28 �C for
2–3 h, and pollen tubes continued to elongate after 8 h of
incubation, indicative of the vitality of pollen (Fig. 3d).
Immersing stigmas in peroxide solution has been used
to measure the stigma receptiveness (Bredemeijer 1982).
In this experiment, the pistils were dissected from 24
flowers just close to anthesis and immersed into a 3 %
H2O2 solution for several minutes. Many oxygen bubbles
were formed and released from the stigma due to the pre-
sence of the peroxidase enzyme (Fig. 3e), indicating the
receptivity of the stigma.
To investigate the growth of pollen tubes in situ, flowers
close to anthesis were either directly collected or bagged in
Cellophane for further 2 days. The pistils were dissected
and stained with aniline blue. Under the fluorescent
microscope, a great number of pollen grains were found to
adhere to the stigmas of the flowers just close to anthesis
and the pollen tubes began to germinate (data not shown).
The pollen tubes were readily visualized on the stigmas of
the bagged flowers (Fig. 3f).
We further tested the self-fertilization of C. pumila
flowers by bagging experiment. The seed setting percent-
age was counted 2 weeks later. Of 60 flowers analyzed, 58
yielded fertile capsules (96.7 % of seed setting percentage;
see Supplementary Table 1). To examine whether C. pu-
mila could be cross-pollinated when flowers open, we
artificially emasculated seven immature flowers of about
1.5 cm in length (mature flowers are 3–4 cm in length),
and then artificially pollinated six flowers 48–72 h later
with one flower served as a negative control. About
2 weeks later, six hand-pollinated flowers all produced
Fig. 3 C. pumila flowers are self-pollinated. a–c A flower just close
to anthesis was dissected longitudinally to show its mature stamens.
Bars, 0.5 cm. d The pollen tubes germinated on culture medium.
Bar, 100 lm. e Activity analysis of the stigma of a flower just before
anthesis. Bar, 1 cm. f Germinated pollen tubes on the stigma of a
bagged flower. Bar, 200 lm
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fertile capsules, whereas the artificially emasculated but
non-pollinated flower failed to fruit (Supplementary
Fig. 1), indicating that C. pumila flowers have a potential
of cross-pollination, facilitating the genetic cross.
Karyotype and genome size analyses of C. pumila
The chromosome number and size of C. pumila were
evaluated by observing root tip cells at the mitotic meta
phase under a microscope (Fig. 4a, b). The result showed that
C. pumila has eight chromosomes of 3.5–6.0 lm in length
(Fig. 4c, d). The karyotype is formulated as 2n = 8 =
6 m ? 2 sm (2 sat) with three pairs of m-chromosomes and
one pair of sm-chromosomes (Fig. 4d; Supplementary
Table 2). The first pair of chromosomes has a secondary
constriction in the interstitial region of both the long and
short arms (Fig. 4d, No. 1 and 2), whereas the second one has
a secondary constriction only in the long arm (Fig. 4d, No. 3
and 4). Two sm-chromosomes have a satellite in the terminal
region of the short arm (Fig. 4d, No. 3 and 4).
We further measured the absolute nuclear DNA amount
(genome size) of C. pumila using flow cytometry. Three
different C. pumila plants were analyzed with rice (Oryza
sativa L. var. Nipponbare) served as an internal reference
standard. According to Burr (2002), the 2C DNA amount of
rice is 0.9 pg. In this experiment, the mean ratio of G1 peaks
(C. pumila : rice) was 1.8 (Supplementary Fig. 2). Therefore,
the 2C DNA amount of C. pumila was estimated to be 1.6 pg.
According to the formula 1 pg DNA = 0.978 9 109 bp
(Dolezel et al. 2007), the haploid genome size of C. pumila
was about 798.7 Mb (Supplementary Table 3).
Shoot induction and antibiotic sensitivity experiments
BA and NAA at different concentrations were added into MS
medium to assess their effects on shoot induction rate. Leaf
explants thickened after about 1 week of culture. About
2 weeks later, adventitious shoots began to appear on the
wound edges of the explants. While MS medium lacking
growth regulators led to a relatively low shoot induction rate
(67.8 ± 10.0 %), the addition of appropriate concentration
of BA and NAA enhanced it. The highest shoot induction rate
(97.6 ± 4.1 %) was obtained when 0.5 mg l-1 BA and
0.1 mg l-1 NAA were applied (Table 1). Under this condi-
tion, abundant adventitious shoots could be induced from
explants within 4 weeks with no appearance of undesirable
roots (Table 1; Supplementary Fig. 3). The result also
showed that relatively lower BA and higher NAA led to high
root induction rate, and it reached up to 95.1 ± 4.3 % when
1.0 mg l-1 NAA were applied (Table 1; Supplementary
Fig. 3). Nevertheless, root induction in C. pumila requires
neither BA nor NAA, and the adventitious shoots could
naturally generate roots after transferring to fresh MS med-
ium without any growth regulator (data not shown). There-
fore, 0.5 mg l-1 BA and 0.1 mg l-1 NAA were applied in
following shoot induction experiments, and MS medium
without any growth regulator was used for root induction.
It is reported that abundant adventitious shoots could be
induced from two Gesneriaceae plants at 25 �C (Tang et al.
2007a, b). Here, to evaluate whether different temperatures
affect the shoot induction rate, fresh leaf explants were
cultured on MS medium supplied with 0.5 mg l-1 BA and
0.1 mg l-1 NAA at different temperatures. The highest
shoot induction rate (97.8 ± 11.6 %) was obtained at
26 �C. Both higher and lower temperatures reduced the
shoot induction rate, and it dropped to 55.5 % at 22 �C
(Supplementary Table 4). However, Fisher’s LSD test
(P \ 0.05) showed that the shoot induction rates obtained
at 24, 26 and 28 �C were not significantly different from
each other, indicating that C. pumila can adapt to a rela-
tively wide temperature range. Nevertheless, for unifor-
mity, an intermediate temperature, i.e. 26 �C was applied
in following experiments.
Hygromycin at different concentrations was added into
MS medium containing 0.5 mg l-1 BA and 0.1 mg l-1
NAA to determine whether it is effective for selecting C.
pumila transformants. The induction of adventitious shoots
was severely affected by hygromycin at the concentration
of 10 mg l-1 or higher. When its concentration reached
to 20 mg l-1, all explants became necrotic with no shoot
Fig. 4 The karyotype analyses of C. pumila. a Resting nucleus.
Bar, 5 lm. b Mitotic prophase chromosomes. Bar, 5 lm. c The eight
chromosomes at the mitosis metaphase. Bar, 5 lm. d The karyotype
of C. pumila. Bar, 5 lm
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induced (Supplementary Table 5). Therefore, 20 mg l-1
hygromycin was used in following transformation
experiments.
Factors affecting hygromycin-resistant shoot induction
frequency in transformation experiments
Several factors affect Agrobacterium inoculation and shoot
induction, including Agrobacterium cell density, infection
time, pre-culture time and co-culture duration (Mondal et al.
2001; Kim et al. 2004; Barik et al. 2005; Crane et al. 2006; Du
and Pijut 2009; Jian et al. 2009). In this study, co-culture time
was first examined by infecting fresh leaf explants with
Agrobacterium LBA4404 of OD600 = 0.6 for 20 min and
culturing on the co-culture medium for 1–5 days. The
highest shoot induction rate (42.2 ± 5.1 %) was achieved
after 2-days of co-culture, and it declined with shortened or
prolonged co-culture (Fig. 5a). 1- and 5-days of co-culture
resulted in the lowest shoot induction rate. Hence, co-culture
for 2 days was applied to next transformation experiments.
To evaluate whether pre-culture could enhance the induc-
tion frequency, newly prepared leaf explants were pre-cul-
tured on MS medium for 0–4 days, inoculated with
Agrobacterium of OD600 = 0.6 for 20 min, and co-cultured
for 2 days. While fresh explants gave rise to the lowest shoot
induction rate (41.7 ± 3.6 %), pre-culture significantly
enhanced it (Fig. 5b). 1-day of pre-culture witnessed the
highest shoot induction rate (95.8 ± 7.2 %), whereas exten-
ded duration led to slightly lower rate. Even though Fisher’s
LSD test (P \ 0.05) showed that 1, 2, 3 or 4 days of pre-
culture resulted in no significant difference in the shoot
induction rate, a pre-culture of 1 day was applied in following
assays due to timesaving and high shoot induction rate.
To analyze whether the OD600 value of Agrobacterium
influences the induction frequency, 1-day pre-cultured leaf
explants were inoculated with Agrobacterium LBA4404 of
different OD600 values for 20 min and co-cultured for
2 days. The highest regeneration frequency (90.0 ± 3.2 %)
was achieved at the late-log phase (corresponding to
OD600 = 0.6), whereas both lower and higher OD600 val-
ues significantly reduced it (Fig. 5c). When OD600 reached
up to 1.0, the lowest regeneration efficiency (52.6 ±
2.8 %) was obtained because of the uncontrollable over-
growth of Agrobacterium.
To verify if Agrobacterium inoculation time affects the
shoot induction rate, 1-day pre-cultured leaf explants were
immersed in Agrobacterium LBA4404 solutions
(OD600 = 0.6) for 10–50 min. 20 min of inoculation was
found to achieve the highest shoot induction frequency
(92.5 ± 5.0 %) (Fig. 5d). It markedly decreased with
increased inoculation time, dropping to 30.0 ± 1.0 %
when the infection time was 50 min because of the over-
growth of Agrobacterium. Although the difference between
20 and 30 min of inoculation was not significant, a rela-
tively short infection period is probably more beneficial for
the viability of explants. Therefore, 20 min of inoculation
was used in following experiments.
GUS activity assays
57 Hyg-resistant plantlets from different leaf explants were
obtained using the optimal conditions, i.e. inoculating
Fig. 5 Effects of different
factors on the shoot induction
rate of C. pumila. Effects of co-
culture time (a), pre-culture
time (b), OD600 value (c), and
infection time (d) on the shoot
induction rate were determined.
Means of induction efficiencies
were compared using a Fisher’s
LSD test (P \ 0.05) and column
bars labeled with the same
letters are not significantly
different. Data, presented as
mean ± SD, were calculated
from three independent
experiments with about 40 leaf
explants each
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1-day pre-cultured explants with Agrobacterium LBA4404
(harboring pCAMBIA1301) for 20 min, culturing on
the co-culture medium for 2 days and selecting on the
selection medium containing 20 mg l-1 hygromycin.
Hygromycin-resistant shoots appeared on the wound edges
of explants after about 4 weeks of induction (Fig. 6a).
After further 4 weeks, the shoots of about 0.5 cm in length
were excised and transferred onto fresh MS medium
Fig. 6 Transgenic C. pumila plants and GUS activity assays.
a Adventitious shoots appeared on the wound edges of leaf explants.
The photo was taken 4 weeks after Agrobacterium inoculation. Bar,
1 cm. b Hygromycin-resistant shoots were transferred onto fresh MS
medium to promote rooting and shoot elongation. Bar, 1 cm. c Shoots
with obvious roots after maintained on MS medium for about 1 week.
Bar, 1 cm. d A transgenic plant (photographed 1 month after
transplantation). Bar, 1 cm. e PCR identification of transgenic plants.
M, 2 kb DNA marker; N, negative control; P, plasmid DNA; 1–7,
seven independent transgenic plants. f GUS staining of wild-type
(left) and transgenic (right) leaves. Bar, 1 mm. g GUS staining of
wild-type (left) and T0 transgenic (right) stems. Bar, 1 mm. h GUS
staining of wild-type (lane 1 and 2) and T1 transgenic (lane 3–6)
leaves
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(without any growth regulator) to promote rhizogenesis and
shoot elongation (Fig. 6b). About 1 week later, the roots
could be readily observed (Fig. 6c). Plantlets of 1–2 cm in
length with well-developed roots were transplanted into
pots. Transgenic plants grew well in culture room with
purple spots appearing slowly on the old leaves (Fig. 6d).
Furthermore, the transgenic plants overexpressing the GUS
gene were morphologically normal comparing with wild-
type ones (Supplementary Fig. 4).
Polymerase chain reaction was carried out to investigate
whether the GUS gene had been introduced into C. pumila.
As a result, specific gene products of expected size
(1,081 bp) were amplified from seven independent trans-
genic plants with no fragment amplified from wild-type
ones (Fig. 6e). GUS activity assays were conducted to
investigate the GUS expression. Both leaves and stems of
transgenic plants were analyzed with wild-type ones served
as negative controls to eliminate the possibility of endog-
enous GUS expression. The results showed that strong and
uniform GUS signal was observed in both leaves and stems
of transgenic plants with no expression signal in wild-type
ones (Fig. 6f, g).
Progenies of line 2 (Fig. 6e) were further analyzed to
validate the GUS gene inheritance by culturing on MS
selection medium containing 25 mg l-1 hygromycin. Of 33
progenies analyzed, 24 could germinate and develop nor-
mally, while the remainder slowly became withered after
germination, conforming to a Mendelian segregation ratio
(3:1) for monogenic inheritance. Subsequent GUS activity
assays showed that these hygromycin-resistant seedlings
could generate GUS signal with no expression signal in
wide-type ones. The above results clearly indicated that the
GUS gene had been introduced into the host and obtained a
stable and uniform expression.
Validation of the transformation system by transferring
the GsNST1B gene
GsNST1B controlling the secondary wall biosynthesis in G.
soja (Dong et al. 2013) was introduced into C. pumila to
validate the availability of this transformation system in
gene function investigation. Positive transgenic plantlets
were confirmed by PCR followed by DNA sequencing to
avoid the possible amplification of endogenous NST-like
genes. The results showed that of 199 hygromycin-resistant
plantlets induced from 55 different leaf explants (for each
explant, 3–4 plants were analyzed by PCR), 23 belonging to
6 different transgenic lines were confirmed to be positive. In
contrast to wild-type plants that had normally developed
leaves (Fig. 7a), transgenic plants showed upward curling
leaves (Fig. 7b–d). Sections of the transgenic and wild-type
leaves were stained with toluidine blue to understand the
cellular basis of upward curling leaves. In wild-type leaves,
the secondarily thickened cells were only found in the veins
and xylem strands (Fig. 7e). In contrast, the parenchyma
mesophyll cells, in addition to the veins and xylem strand
cells were heavily secondarily thickened in the transgenic
leaves (Fig. 7f). RT-PCR was further carried out to check
the expression of the GsNST1B gene in transgenic plants. As
shown in Fig. 7g, the GsNST1B gene was strongly expressed
in three independent transgenic plants with no signal in
wild-type ones. In addition, GsNST1B overexpressors gen-
erated undeveloped fruits (data not shown), similar to our
previous report (Dong et al. 2013).
Discussion
Characterization of C. pumila as an ideal model plant
for evo-devo studies
The haploid chromosome number n = 4 of C. pumila was
previously reported by Ratter (1963). We here document its
karyotype 2n = 8 = 6 m ? 2sm (2sat), the lowest number
of chromosomes reported in Gesneriaceae to date (Skog
1984; Li and Wang 2004; Weber 2004; this study). The
species in Gesneriaceae are usually polyploidy with
perennial habit (Skog 1984; Li and Wang 2004; Weber
2004). The genus Chirita sensu stricto is one of the rarely
occurred diploid taxa with annual habit in Gesneriaceae
(the traditional polyphyletic Chirita was split into four
monophyletic groups including the perennial Primulina
and Liebigia and the annual Chirita sensu stricto and
Microchirita; see Wang et al. 2011). The close relatives of
C. pumila in Chirita usually have diploid chromosome
number of 2n = 18 (Li and Wang 2004). Given that the
basic chromosome number is x = 9 or 8 in Gesneriaceae,
the low chromosome number of C. pumila was assumed to
be achieved through successive unequal translocation from
a complement of chromosomes with x = 9, probably cor-
related with its short-lived habit (Ratter 1963; Skog 1984;
Li and Wang 2004). Researches addressing chromosome
evolution between C. pumila and its relatives would reveal
how the rearranged chromosomes contribute to plant habit
shifts, reproductive isolation and speciation.
In addition, we found a special phenomenon that C.
pumila flowers perform self-fertilization before anthesis,
i.e. cleistogamy, an extreme form of self-fertilization first
reported in Gesneriaceae. Selfing has commonly been
viewed as an ‘‘evolutionary dead end’’ because it usually
leads to inbreeding depression by accumulating recessive
deleterious alleles (Stebbins 1957; Barrett 2002; Boggs
et al. 2009). However, when environments become fluc-
tuant and unpredictable with scarcity or inconsistency of
pollinators or population bottlenecks, selfing rates would
increase with inbreeding depression gradually overcome
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because deleterious alleles may be purged over successive
generations of selfing (Barrett 2002; Boggs et al. 2009;
Albert et al. 2011). The self-fertilization will finally
become established owing to adaptive advantages of self-
pollination in providing reproductive assurance when out-
crossing fails (Darwin 1876). Morphologically, the con-
version from outcrossing to cleistogamy involves the shifts
from dichogamy to homogamy and from herkogamy to
plesiogamy, and precocious maturation of sexual organs
(Campbell et al. 1983). In Gesneriaceae, almost all species
have hermaphroditic flowers containing both female and
male sexual organs which usually spatially separate in a
flower, i.e. herkogamy for cross-pollination. Sexually
mature pollen and stigmas are presented as flowers bloom
to attract animal pollinators (Wang et al. 2010, 2011). In
the evolutionary transition from cross-fertilization to
cleistogamy, a series of floral morphological and physio-
logical modifications have occurred in C. pumila, including
the same position of the anthers and stigmas just below the
upper inner surface of the corolla tube, and the precocious
and simultaneous maturation of pollen and stigmas before
anthesis. In addition, the flowers of C. pumila open with
anthers included and stigma exserted. Given hand-
pollinated flowers producing fertile fruits, C. pumila
should have the possibility of cross-fertilization, a mixed
mating system envisaged as a ‘‘bet-hedging strategy’’ for
fluctuating and unpredictable environments (Berg and
Redbo-Torstensson 1998; Culley and Klooster 2007). The
typical cleistogamous flowers with potential cross-polli-
nation make C. pumila an ideal candidate model to
understand the ecological success of natural selection and
genetic mechanisms for the mating systems of cleistogamy
versus chasmogamy.
The ABCE model is a widely used framework to
understand the floral development and evolution in angio-
sperms (Soltis et al. 2007; Litt and Kramer 2010). How-
ever, some components of the ABCE model, such as
A-function floral identity genes, are so far limited to
Arabidopsis and its close relatives and their functions have
not yet been testified in other lineages of angiosperms (e.g.
Antirrhinum; Litt 2007; Bowman et al. 2012). Additional
function analyses in emerging evo-devo model organisms
are therefore critically important to finally elucidate whe-
ther the BC model lacking the A-function is general in
eudicots (Litt 2007; Soltis et al. 2007; Causier et al. 2010;
Bowman et al. 2012). In addition, the origin of zygomor-
phic flowers is suggested to be one key innovation asso-
ciated with the explosive radiation of angiosperms (Dilcher
2000; Cubas 2004; Busch and Zachgo 2009). Increasing
evidence indicates that CYC-like TCP genes play a crucial
role in the origin and evolution of floral zygomorphy in
angiosperms (Cubas 2004; Busch and Zachgo 2009;
Fig. 7 Analysis of transgenic plants overexpressing GsNST1B. a–dOne wild-type plant (a) and three independent transgenic plants (b–d)with upward curling leaves. Bars, 1 cm. e Toluidine blue staining
results showing normal secondary wall thickening of wild-type
plants in the veins and vascular bundles. Bar, 10 lm. f The ectopic
secondary wall thickening of transgenic plants in parenchyma
mesophyll cells. Bar, 10 lm. g RT-PCR analysis of GsNST1B in
three independent transgenic plants with wild-type plant served as a
negative control. CpACTIN was amplified as an internal reference
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Preston and Hileman 2009; Specht and Bartlett 2009). A
recent report suggests that the repeated origins of floral
zygomorphy are related to the independent gains of similar
positive autoregulatory elements in CYC-like TCP genes in
different lineages of angiosperms (Yang et al. 2012).
However, it is still a challenge to decipher how these genes’
activities are controlled by upstream factors, including the
dorsal identity function, the functional domain expansion to
lateral or ventral floral regions, the loss-of-function and so
on (Song et al. 2009; Martın-Trillo and Cubas 2010; Yang
et al. 2012; Hileman 2014). C. pumila is apparently an ideal
model species to address these questions because of its
phylogenetic representativeness, annual habit, short life
cycle, and self-fertility and diploid, the widely accepted
selection criteria for evo-devo model organisms (Irish and
Benfey 2004; Jenner 2006; Jenner and Wills 2007; Sommer
2009; Ankeny and Leonelli 2011). Its diploid with low
chromosome number would facilitate the identification of
recessive traits and avoid the complication of gene dosages,
and its typical cleistogamous flowers with potential cross-
pollination enable C. pumila to be maintained with homo-
zygous lines straightforward and capable of generating
genetic crosses. Therefore, we here select C. pumila as a
target to develop the genetic transformation system.
Efficient Agrobacterium-mediated transformation
and regeneration of C. pumila
An efficient Agrobacterium-mediated transformation and
regeneration system is developed in C. pumila in this study,
which depends on a powerful shoot induction ability of
leaf explants on MS medium containing 0.5 mg l-1 BA
and 0.1 mg l-1 NAA (Table 1). Moreover, similar to
Perilla frutescens (Kim et al. 2004), the adventitious shoots
of C. pumila directly form on the wound edges of leaf
explants without an evident callus phase. In addition, C.
pumila has a powerful rooting ability because no growth
regulator is required and roots can be readily observed after
1 week of culture on fresh MS medium. Therefore, C.
pumila has a powerful regeneration ability using leaf
explants, a prerequisite for genetic transformation.
High efficient gene transfer and powerful regeneration
ability of explants after Agrobacterium inoculation are
crucial to plant transformation (He et al. 2010). Many
factors may affect the transformation frequency and
thereafter the shoot induction rate to varying degrees. As a
pivotal step in transformation process, co-culture of the
inoculated explants with Agrobacterium allows T-DNA
transfer from plasmid into plant cells. In general, co-culture
for 2–3 days reaches the highest transformation frequency
(Kim et al. 2004; Jian et al. 2009). However, co-culture
duration can be prolonged to 4–5 days for some species
(Mondal et al. 2001; Barik et al. 2005). Here, 2 days of
co-culture reaches the highest shoot induction rate. While
1 day of co-culture is insufficient for Agrobacterium
infection and T-DNA transfer, extended duration may
cause the damage of explants owing to the uncontrollable
overgrowth of Agrobacterium. Accordingly, the shoot
induction frequency is reduced in both cases.
It is reported that pre-culture of explants prior to
Agrobacterium inoculation can significantly enhance the
transformation frequency in Cajanus cajan (Lawrence and
Koundal 2000), Lathyrus sativus (Barik et al. 2005), Lotus
corniculatus (Jian et al. 2009) and Fagopyrum esculentum
(Chen et al. 2008). However, pre-culture drastically
declines the transformation competence of Citrus paradise
(Costa et al. 2002) and Perilla frutescens (Kim et al. 2004).
In this study, pre-culture enhances drastically the shoot
induction rate, indicating a positive effect of pre-culture on
C. pumila transformation probably due to the improved
viability of explants. However, extended pre-culture redu-
ces slightly the shoot induction rate probably by dimin-
ishing the susceptibility of explants to Agrobacterium,
indicating that only appropriate pre-culture duration is
benefit for the transformation and regeneration of
C. pumila.
Agrobacterium cell density and inoculation time can
also affect transformation efficiency (Du and Pijut 2009;
Jian et al. 2009). While low Agrobacterium cell concen-
tration and short infection time may result in insufficient
attachment of Agrobacterium to explants and reduce the
transformation frequency, increased Agrobacterium cells
and prolonged inoculation time would damage explants
and decrease the regeneration frequency. Here, the highest
shoot induction frequency is obtained when fresh leaf
explants are infected with Agrobacterium cells of
OD600 = 0.6 for 20 min. Both high and low Agrobacte-
rium cell density, as well as shortened or prolonged inoc-
ulation time reduce the shoot induction rate, indicating that
appropriate Agrobacterium cell density and inoculation
time are important for successful transformation.
As outlined above, a high shoot induction frequency is
achieved in C. pumila by infecting 1-day pre-cultured
explants with Agrobacterium of OD600 = 0.6 for 20 min
followed by 2-days of co-culture. Further PCR and GUS
activity assays of T0 and T1 plants indicate that the GUS
gene has been successfully introduced into the host, evident
by its stable and uniform expression. However, further
transformation experiment using a gene with known
function and obvious phenotypic effects is required to
validate the applicability of a transformation system in
gene function investigation (Jian et al. 2009). Here, the
GsNST1B gene implicated in secondary wall biosynthesis
in G. soja is selected due to its obvious phenotype during
early vegetative growth stages in Arabidopsis overexpres-
sors (Dong et al. 2013). Similar to Dong et al. (2013),
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GsNST1B overexpression in C. pumila generates a desired
phenotype characteristic of upward curling rosette leaves
that is attributed to the ectopic thickening of secondary
walls, indicating the applicability of this transformation
system in gene function investigation.
A reliable and efficient transformation system is crucial to
comparative functional studies in evo-devo that aims at
exploring the evolutionary mechanisms underlying morpho-
logical changes. However, developing a transgenic system is
usually time-consuming and laborious because it requires
several generations of subculture and alterations of medium.
In Lotus japonicas and Medicago truncatula, for example,
about 4 months are needed to produce transgenic plants
(Stiller et al. 1997; Crane et al. 2006). In Lotus corniculatus,
the superroot- derived transformation protocol is complicated
with at least five changes of medium (Jian et al. 2009).
In Triticum turgidum, obtaining transgenic lines requires 2–3
rounds of selection (He et al. 2010). As a classical model plant,
Antirrhinum has been proved to be successful in Agrobacte-
rium-mediated genetic transformation accompanied by
repeatedly improved transformation protocol (Cui et al. 2003,
2004; Manchado-Rojo et al. 2012). Nevertheless, it still leaves
something to be desired that might restrict its wide applica-
tion. In this study, using leaf disks as explants, the transfor-
mation process (from Agrobacterium inoculation to PCR
identification) takes about 3 months with a high efficiency
(Fig. 8). In addition, the entire transformation process is
simple because no specific rooting media is required and the
shoot induction and selection are achieved in one step. Taken
together, the C. pumila transformation system has the features
of simplicity, rapidity and high-efficiency.
Since its inception at the end of last century, evo-devo
has passed from an initial stage to a rapid developing
discipline, evident by emerging model organisms in both
animals and plants. With completion of genome sequenc-
ing project, establishment of a mutant library and further
optimization of the transformation system, C. pumila could
become a fascinating model plant for a wide range of evo-
devo studies, especially in the field of floral symmetry,
floral organ identity, chromosome evolution and mating
system evolution.
Acknowledgments We thank James F. Smith for his constructive
comments and language improvements on this article. This work was
supported by the National Natural Science Foundation of China
(30990240 and 31170198).
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Fig. 8 A flowchart for Agrobacterium-mediated transformation of
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